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C H A P T E R
T W E LV E
Controlled Expression of
Iron-Sulfur Cluster Assembly
Components for Respiratory Chain
Complexes in Mammalian Cells
Oliver Stehling, Alex D. Sheftel, and Roland Lill
Contents
1. Introduction
2. Depletion of Fe/S Cluster Assembly Components
by RNA Interference
2.1. Vector-based RNAi
2.2. siRNA design
2.3. Electroporation-based transfections
2.4. Assessing the efficiency of the RNAi treatment
2.5. Assessing the specificity of the RNAi treatment
3. Analysis of Respiratory Complex Assembly
3.1. Assessing the incorporation of iron-containing cofactors into
respiratory complexes
3.2. Assessing the subunit composition of respiratory complexes
by two-dimensional BN-PAGE
4. Analysis of Respiratory Complex Function
4.1. Determination of lactate formation
4.2. Determination of complex I activity by in-gel activity staining
4.3. Determination of complex I activity by spectrophotometry
4.4. Analysis of enzyme activities in multiwell plates
5. Concluding Remarks
Acknowledgments
References
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Abstract
Three of the respiratory chain complexes contain essential iron-sulfur (Fe/S)
cluster prosthetic groups. Besides respiration, these ancient inorganic cofactors
are also necessary for numerous other fundamental biochemical processes in
Institut für Zytobiologie and Zytopathologie, Philipps-Universität, Marburg, Germany
Methods in Enzymology, Volume 456
ISSN 0076-6879, DOI: 10.1016/S0076-6879(08)04412-1
#
2009 Elsevier Inc.
All rights reserved.
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virtually every known organism. Both the synthesis of Fe/S clusters and their
delivery to apoproteins depend on the concerted function of specialized, often
dedicated, proteins located in the mitochondria and cytosol of eukaryotes.
Impaired function of the mitochondria-located Fe/S cluster (ISC) assembly
machinery affects all cellular Fe/S proteins, including enzymes of the respiratory chain, NADH: ubiquinone oxidoreductase (complex I; eight Fe/S clusters),
succinate: ubiquinone oxidoreductase (complex II; three Fe/S clusters), and
cytochrome bc1 complex (complex III; one Fe/S cluster). Here, we describe
strategies and techniques both to deprive respiratory chain proteins of their
Fe/S cofactors and to study changes in activity and composition of these
proteins. As examples, we present the results of the depletion of two types of
Fe/S biogenesis proteins, huNfs1 and huInd1, in a human tissue culture model.
The ISC assembly component huNfs1 is required for biogenesis of all cellular
Fe/S proteins, its loss exerting pleiotropic effects, whereas huInd1 is specific
for Fe/S cluster maturation of complex I. Disorders in Fe/S cluster assembly are
candidate causes for defects in respiratory complex assembly of unknown
etiology.
1. Introduction
The respiratory chain of mammalian mitochondria is a multienzyme
system comprising more than 80 polypeptide chains, assembled into five
individual complexes at the mitochondrial inner membrane (Scheffler,
2007). Together with two electron carriers, coenzyme Q and cytochrome c,
complexes I to IV constitute an electron transport chain, transferring electrons
from NADH and FADH2 to molecular oxygen. This electron flow results in a
concomitant translocation of protons across the inner membrane into the
intermembrane space, thereby establishing a proton motive force that finally
drives complex V to generate ATP.
Electron transfer within the electron transport chain is conducted by
low-molecular mass, serially arranged redox components, including flavins,
Fe/S clusters, quinones, heme species, and copper centers. Consequently,
the formation of each of the multi subunit respiratory complexes not only
requires the concerted action of numerous protein assembly factors
(Coenen et al., 2001) but also depends on the functionality of many systems
involved in cofactor biosynthesis and insertion. The most abundant redox
centers within the electron transport chain are Fe/S clusters located in
respiratory complexes I, II, and III. So far, three different biogenesis systems
required for the de novo synthesis of Fe/S clusters and their incorporation
into apoproteins have been identified in nongreen eukaryotes: the ironsulfur cluster (ISC) assembly and the ISC export machineries located within
mitochondria, and the cytosolic Fe/S protein assembly (CIA) apparatus
[for a recent comprehensive review see Lill and Mühlenhoff (2008)].
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Controlled Expression of Fe/S Assembly Components
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Among these systems, the mitochondrial ISC assembly machinery holds a
central role in that it is required for the biogenesis of all cellular Fe/S
proteins. One of the key events catalyzed by this machinery is the abstraction of sulfur from cysteine by a heteromultimeric complex consisting of
the pyridoxal 50 -phosphate (PLP)–dependent cysteine desulfurase Nfs1 and
the eukaryote-specific component Isd11 (Adam et al., 2006; Lill and
Mühlenhoff, 2008; Wiedemann et al., 2006). During catalysis, the cysteine
substrate is transiently bound to the PLP cofactor of Nfs1, and the substrate
cysteine sulfur is then attacked by an active-site cysteinyl residue to produce
a ‘‘persulfidic sulfur’’ (Zheng et al., 1993, 1994). The activated sulfur is
subsequently available for the formation of a transient Fe/S cluster on the
scaffold protein Isu1. This step involves the assistance of frataxin as an iron
donor and of the electron transfer chain NADH—ferredoxin reductase—
ferredoxin possibly required for reduction of the sulfur to sulfide (Lill and
Mühlenhoff, 2008). Finally, the Isu1-bound Fe/S cluster is delivered to
recipient apoproteins, a process aided by a dedicated Hsp70 chaperone
system and the monothiol glutaredoxin Grx5 (Craig et al., 2006; Herrero
and de la Torre-Ruiz, 2007).
Fe/S protein assembly within mitochondria is connected to the respective extramitochondrial process by the ISC export machinery. The central
component of this latter system is an ABC transporter of the mitochondrial
inner membrane, termed Atm1 in yeast and ABCB7 in mammals, which
appears to export a still unknown compound to the cytosol, where it is used
by the CIA components for maturation of extramitochondrial Fe/S proteins
(Cavadini et al., 2007; Kispal et al., 1997; Pondarré et al., 2006). According
to a current model, the cytosolic P-loop NTPases Nbp35 and Cfd1 (in
mammals also known as Nubp1 and Nubp2, respectively) form a heterotetrameric complex and serve as a scaffold to assemble a transient Fe/S
cluster (Lill and Mühlenhoff, 2008; Netz et al., 2007; Stehling et al.,
2008). The labile metallocluster is then transferred to target apoproteins, a
process that has been shown in yeast to require the function of the iron-only
hydrogenase-like protein Nar1 (in mammals known as IOP1) and the
WD40 repeat protein Cia1 (termed Ciao1 in mammals) (Lill and
Mühlenhoff, 2008; Song and Lee, 2008).
Because key components of the mitochondrial ISC assembly machinery
participate in the maturation of respiratory complexes I, II, and III by means
of formation of their respective Fe/S clusters (Biederbick et al., 2006; Fosset
et al., 2006; Lill and Mühlenhoff, 2008; Puccio et al., 2001; Rötig et al.,
1997; Song and Lee, 2008; Stehling et al., 2004), these components can be
referred to as general assembly factors for these enzymes. In addition, a
specific assembly component termed Ind1 (iron-sulfur protein required for
NADH-dehydrogenase) has been identified that assists in the formation
of respiratory complex I (NADH: ubiquinone oxidoreductase, NADH
reductase), the largest enzyme of the mammalian respiratory chain
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(Bych et al., 2008; Sheftel et al., manuscript in preparation). Complex I
consists of 45 subunits arranged in an L-shaped manner with one arm
embedded into the inner membrane and the other one protruding into
the mitochondrial matrix (Brandt, 2006; Vogel et al., 2007). The latter arm
harbors a noncovalently bound FMN cofactor involved in NADH oxidation and eight Fe/S clusters (Hinchliffe and Sazanov, 2005). Seven of them
form an ‘‘electrical wire’’ to bridge the distance between the catalytically
active FMN center to the ubiquinone reduction site (Brandt, 2006). Until
now, only a few assembly factors primarily required for the maturation of
complex I have been identified. Three of them, NDUFAF1, B17.2L, and
Ecsit, are aiding the assembly of complex I subcomplexes (Vogel et al.,
2007). The fourth, the aforementioned Ind1, was recently identified in both
the yeast Yarrowia lipolytica and in human cells as a mitochondrial homolog
of the cytosolic CIA components Nbp35 and Cfd1 that facilitates the
assembly of Fe/S clusters and subunits of complex I (Bych et al., 2008;
Sheftel et al., manuscript in preparation).
In this chapter, we describe approaches to analyze the role of human Nfs1
(huNfs1) as a general ISC assembly component and of human Ind1 (huInd1)
as a specific assembly factor for respiratory complex I. As basic strategies, we
first deplete the assembly proteins of interest by RNAi technology in tissue
culture cells and then analyze the loss-of-function phenotypes by Fe/S
enzyme activity measurements, 1-D and 2-D gel electrophoresis, and the
incorporation of 55Fe into Fe/S proteins. The experimental techniques
described in the following are generally applicable to study the function of
Fe/S protein assembly factors, and, in particular, may be useful in future
analyses of Fe/S cluster assembly into respiratory complexes.
2. Depletion of Fe/S Cluster Assembly
Components by RNA Interference
2.1. Vector-based RNAi
RNA interference (RNAi) is a mechanism by which small interfering RNAs
(siRNAs) drive an endogenous machinery to degrade distinct RNA target
molecules (Wu and Belasco, 2008). Although duplex siRNAs can be directly
administered to cell culture cells, we have good experience with the vectorbased endogenous production of siRNAs for inducing the degradation of a
specific mRNA (Sandy et al., 2005). Usually, we use the pSUPER vector
developed by Brummelkamp et al. (2002) or one of its commercially available derivates (OligoEngine, Seattle, WA, USA). The vector contains a
polymerase-III H1-RNA gene promoter that drives the expression of socalled short hairpin RNAs (shRNA) that are intracellularly processed to
siRNA duplices of 19 nucleotides in length.
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2.2. siRNA design
Because only guidelines but no unequivocal rules for the selection of
appropriate siRNA sequences exist (Birmingham et al., 2007), we usually
test three different mRNA target sites and choose the ones with the
strongest effect on the intended protein. Tools for the design of siRNAs
are available at numerous websites (a repertory is provided by Pei and
Tuschl, 2006). Alternately, the choice of useful siRNAs may be facilitated
by testing commercially available sets of siRNAs that are selected by
computational methods performed by the manufacturers. In the case of
huNfs1 and huInd1, we finally opted for the 19mer gene-specific targeting
sequences GCACCATTATCCCGGCTGT (positions 1041 to 1059 of the
huNfs1 coding region) (Biederbick et al., 2006) and GCAGAAACCGATAGAAGGT (positions 177 to 195 of the huInd1 coding region) (Sheftel
et al., manuscript in preparation). On the basis of these sequences, 64mer
oligonucleotides were synthesized (Fig. 12.1A) consisting of a BglII restriction site-compatible nucleotide quadruplet, a cytosine triplet, the mRNA
target sequence, a nonameric spacer encoding the shRNA hairpin
(TTCAAGAGA), the mRNA target sequence in its antisense orientation,
a thymidine quintuplet to terminate shRNA expression, and a terminal
GGAAA sequence (Brummelkamp et al., 2002). Corresponding complementary oligonucleotides are designed to carry a HindIII-compatible nucleotide quadruplet at their 50 end but to lack the BglII-compatible sequence at
the 30 end. After annealing and phosphorylation, the resulting oligomeric
DNA duplexes are cloned into the pSUPER vector by means of its BglII/
HindIII restriction sites according to standard procedures, yielding the
RNAi vectors huNFS1-R3 and siIND1, respectively.
2.3. Electroporation-based transfections
Because HeLa cells (human cervix carcinoma) are easy to handle and
transfect, we routinely use this model system to study the maturation of
respiratory chain complexes (Biederbick et al., 2006; Sheftel et al., manuscript in preparation; Stehling et al., 2004). To introduce plasmids and/or
siRNA duplexes, cells are transfected by electroporation. After harvesting
by trypsinization and washing in transfection buffer (21 mM HEPES,
137 mM NaCl, 5 mM KCl, 0.7 mM Na2HPO4, and 6 mM dextrose, pH
7.2) (Chu et al., 1987), typically approximately 4 106 cells are resuspended
in 525 ml transfection buffer and supplemented with 25 mg of the required
plasmids or with 15 mg of the respective siRNA duplexes. The deployment
of up to twice as many cells is possible but may affect transfection efficiency. Cells are transferred into a 4-mm-gap electroporation cuvette and
immediately transfected to prevent sedimentation within the cuvette. Electroporation is carried out at room temperature with an EASYJectþ device
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A
huNfs1
targeting sequence
Bg/II
huNfs1
targeting sequence
(inverse repeat)
Spacer
Terminator
5 - gatc CCC GCACCATTATCCCGGCTGT TTCAAGAGA ACAGCCGGGATAATGGTGC TTTTT GGAAA - 3
5 - tcga TTTCC AAAAA GCACCATTATCCCGGCTGT TCTCTTGAA ACAGCCGGGATAATGGTGC GGG - 3
HindII
B
-R3
-R3
PER uNFS1
pSU
h
PER uNFS1
h
pSU
a-huNfs1
3 days (1st)
6 days (2nd)
Growth time (transfection)
Events
U
SSC
1023
C
U
FSC
1023
pSUPER
siIND1
FL1 (hulnd1-EGFP)
Figure 12.1 RNAi-mediated depletion of huNfs1 and huInd1. (A) Sequences of a
64mer oligonucleotide hairpin insert (top) and of its complementary strand (bottom)
encoding a shRNA species that is directed against huNFS1 mRNA. (B) HeLa cells were
transfected twice by electroporation in 3-day intervals with either the empty pSUPER
vector or the pSUPER-derived vector huNFS1-R3 containing the 64mer oligonucleotide hairpin insert of part (A). Cell lysates obtained 3 days after each round of transfection were immunoblotted and stained for huNfs1. (C) HeLa cells were transfected
by electroporation with a vector encoding a huInd1-EGFP fusion protein either in combination with the empty pSUPER vector or with the huInd1 mRNA-directed siRNA
vector, designated siIND1 (20 mg and 15 mg, respectively). Three days later, EGFP-fluorescence was determined by flow cytometry (fluorescence channel 1, FL1) and served as
a measure for the efficiency of siIND1 to deplete the mRNA of the tagged huInd1 version.The insert shows the physical parameters of huInd1-EGFP^expressing cells and the
region used for gating. FSC, Forward scatter; SSC, side scatter.
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Controlled Expression of Fe/S Assembly Components
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with settings of 250 V and 1500 mF, resulting in a pulse time of approximately 30 msec. Considerably longer pulse times should be avoided,
because they will lead to massive cell damage, whereas shorter pulses will
decrease the transfection efficiency. Immediately after electroporation
(essentially within 30 sec, according to the manufacturer’s recommendations), cells are transferred to culture medium supplemented with 20%
conditioned HeLa medium (HeLa tissue culture supernatant) and grown
on an area of 75 cm2. The addition of conditioned medium is strongly
suggested, because it substantially improves recovery and yield of transfected
cells (unpublished observations).
Transfection of the huNFS1-R3 or siIND1 RNAi vector will decrease
the target protein levels nearly to the detection limit as early as 3 to 4 days
after electroporation (Biederbick et al., 2006; Sheftel et al., manuscript in
preparation). Only beginning at this point, can the maturation of Fe/S
subunits of respiratory complexes be assumed to be affected. Hence, the
occurrence of conspicuous cellular phenotypes requires at least another 3 or
4 days. However, because of the transient character of the transfection,
vector-mediated RNAi effects will disappear over time (Stehling et al.,
2004). To prolong the time period of huNfs1 or huInd1 depletion, HeLa
cells have to be retransfected, under conditions identical to the first electroporation. Principally, these retransfections can be carried out every third to
fourth day as long as it is necessary. Time intervals shorter than 3 or longer
than 4 days are not recommended. On the one hand, cells require time
to recover from the electroporation procedure, and on the other hand,
huNfs1 and huInd1 protein levels will start to recuperate (unpublished
observations).
2.4. Assessing the efficiency of the RNAi treatment
Electroporation is an effective but invasive method to transfect cells
(Meldrum et al., 1999; Stehling et al., 2004). One may expect the loss of
one fourth to up to one half of the deployed cells. Transfection efficiency
can be assessed by inclusion of 5 to 10 mg of an EGFP-encoding reporter
vector in the electroporation mixture. Analysis of cell-associated fluorescence 3 to 4 days after the first transfection by flow cytometry usually results
in more than 80% GFP-positive cells. Retransfection may increase the
proportion of transfected cells to more than 90%, leading to the rapid
development of a nearly homogeneous cell population without any need
for selection or lineage generation. Consequently, changes in composition
and activity of the respiratory chain can be monitored rather early after the
depletion of assembly factors, as documented for huNfs1 or huInd1
(Biederbick et al., 2006; Sheftel et al., manuscript in preparation).
Successful and thorough depletion of target proteins is a prerequisite for
analysis of RNAi-mediated loss-of-function phenotypes. Although the
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determination of mRNA levels (e.g., by Northern blotting or quantitative
PCR) encoding the intended target proteins is a useful indicator for the
action of siRNAs, we prefer to directly assess the protein levels of the
investigated assembly factors (Biederbick et al., 2006; Sheftel et al., manuscript in preparation). First, only the depletion of the proteins under study
most reliably reflects the phenotypical consequences. Second, stability and
turnover of the intended proteins are both independent of the RNAi
efficiency. For example, huNfs1 and huInd1 levels are analyzed by reducing
SDS-PAGE followed by immunoblotting and compared with the amounts
of endogenous actin or tubulin. Although after the first round of transfection the levels of RNAi target proteins are usually still above the detection
limit, they frequently drop below after retransfection (Fig. 12.1B). When no
specific antibodies are available, the efficiency of siRNAs to deplete the
intended proteins may be assessed by vector-based coexpression of an
epitope-tagged version of the respective target protein. Depending on the
expression levels, translation of the fusion proteins may not be completely
abrogated by RNAi but will be lowest in case of the most efficient siRNA
species (Fig. 12.1C).
2.5. Assessing the specificity of the RNAi treatment
Although siRNAs are designed to be directed solely to one distinct target,
usually an mRNA, they may nonspecifically affect unintended targets, socalled off-targets, in an unpredictable manner (Birmingham et al., 2006;
Cullen, 2006; Jackson et al., 2006b). Consequently, a strong cellular phenotype caused by an allegedly efficient siRNA is not necessarily a specific
one related to the depletion of the intended target as found for an siRNA
directed against the mRNA of the cytosolic huInd1 homolog huNbp35
(Stehling et al., 2008). Because intrusion into the respiratory chain by RNAi
will affect multiple cellular pathways, global gene expression studies are
neither suitable nor feasible to discriminate between specific and nonspecific
RNAi effects. Instead, two different strategies have been developed to
minimize the risk of off-target effects (Chatterjee-Kishore and Miller,
2005; Cullen, 2006). One strategy suggests the application of a pool of
multiple, often chemically modified, siRNA duplexes ( Jackson et al., 2006a)
to minimize the contribution of one individual siRNA species to the allover
RNAi-induced cellular phenotype. However, because the siRNA duplexes
are frequently not individually tested, the specificity of the total pool can
be estimated only by statistical considerations but evades unequivocal practical evaluation. Consequently, this strategy is only recommendable when
the following second approach is difficult to apply.
Our preferred strategy is to verify the specificity of individual siRNA
duplexes by complementation of the RNAi-mediated cellular phenotypes
on the basis of the expression of RNAi-resistant versions of target proteins.
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Controlled Expression of Fe/S Assembly Components
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For instance, the consequences of huNFS1-R3-mediated huNfs1 deficiency can be specifically remedied by coexpression of murine Nfs1
(muNfs1) (Biederbick et al., 2006). Both the mRNA and protein sequences
of muNfs1 are nearly identical to huNfs1. However, at the siRNA target
site, muNFS1 mRNA differs in four nucleotide residues from the huNFS1
mRNA (Fig. 12.2A). These four mismatches are sufficient to prevent
muNFS1 mRNA from RNAi-mediated degradation and to allow the
heterologous, vector-based expression of muNfs1. Because cellular proteins, especially assembly factors, are part of a coordinated proteinaceous
network, expression levels have to be empirically optimized to achieve best
complementation. Usually, we titrate the amount of the complementing
vector to modify cellular expression levels. As a rule, we apply 2.5, 7.5, and
22.5 mg of the complementing vector to the transfection mixture and
choose the amount of DNA that complements best. On the basis of these
results, a second round of titration with closer increments might sometimes
be useful to further narrow down the optimal amount of DNA.
In an alternate approach, the RNAi-mediated huInd1 deficiency was
complemented by vector-based expression of a mutated huInd1 version
(smIND1) whose mRNA contained seven silent mutations within the
siIND1 target site (Sheftel et al., manuscript in preparation) (Fig. 12.2B).
As few as four point mutations may suffice to confer resistance against
RNAi as found for muNFS1 mRNA. Although mismatches are able to
prevent siRNA-mediated cleavage of mRNA, they may instead lead to
translational stalling (Doench et al., 2003; Saxena et al., 2003; Zeng et al.,
2003). Thus, the more silent mutations are introduced into the siRNA
target site, the less likely RNAi-mediated gene silencing becomes. A
versatile PCR-based method to introduce a whole array of closeby
Figure 12.2 Complementation of RNAi-mediated huNfs1 and huInd1 depletion.
(A) Partial nucleotide sequence alignment of muNFS1, huNFS1, and the huNFS1directed siRNA sequence that is part of RNAi vector huNFS1-R3 (cf. Fig. 12.1A and
12.1B). Mismatches in muNFS1 are underlined in bold. (B) Partial nucleotide sequence
alignment of silently mutated huIND1 (smIND1), huIND1, and of the huIND1-directed
siIND1. Mismatches in smIND1are underlined in bold.The numbers indicate the nucleotide positions of the mRNA sequence starting at the AUG start codon.
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mutations at the same time into a vector has been developed by Zheng et al.
(2004). Once the mutations have been established, optimal complementation has to be determined by appropriate titration of smIND1 expression
levels (see earlier).
3. Analysis of Respiratory Complex Assembly
Depletion of huNfs1 or huInd1 has profound effects on mitochondrial
electron transport complexes (Biederbick et al., 2006; Fosset et al., 2006;
Sheftel et al., manuscript in preparation). A multitude of assays has been
established to analyze the activity of the respiratory chain as a whole or of its
individual complexes. In the following sections we will mainly focus on
assays that we have adapted to directly assess the presence of iron-containing
cofactors including Fe/S clusters in individual respiratory chain complexes
or to determine changes in the function of respiratory complexes at high
performance with a microtiter plate reader device.
3.1. Assessing the incorporation of iron-containing cofactors
into respiratory complexes
Proper maturation of respiratory complexes I to IV includes the appropriate
incorporation of the various cofactors. The presence of iron, in the form of
heme or Fe/S cluster cofactors, within the individual respiratory complexes
can be directly assessed on incorporation of radioactive iron (55Fe) followed
by blue-native polyacrylamide gel electrophoresis (BN-PAGE) (Schägger,
2003) and autoradiography. Because of their high Fe/S cluster content,
respiratory complexes I and II will be radiolabeled at high specificity. Cells
lacking a general ISC assembly factor like Nfs1 will contain less radioactive
iron in both enzymes, whereas cells lacking the specific assembly factor Ind1
will incorporate less 55Fe only in respiratory complex I.
The principal route by which mammalian cells take up iron is by receptormediated endocytosis of transferrin (Tf ), a soluble protein containing two
binding sites for ferric iron (Richardson and Ponka, 1997). To provide HeLa
cells the radioactive isotope 55Fe in a physiologic manner it has to be coupled
to transferrin basically as described by Ponka and Schulman (1985).
3.1.1. Preparation and application of 55Fe-loaded transferrin
Dilute 55FeCl3 (approximately 250 nmol 55Fe3þ, equivalent to an activity of
1 mCi) in 1 ml of 0.1 N HCl and add a 50% (w/v) sodium citrate solution in
a 100-fold molar excess. The color of the mixture will turn from clear to
pale yellow. After an incubation of at least 3 h at room temperature add to
approximately 60 mM apotransferrin dissolved in 0.6 M sodium hydrogen
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carbonate (bicarbonate, NaHCO3) to a final volume of 2 ml. The color of
the mixture will now turn from yellow to salmon. The final molar ratio of
transferrin to 55Fe3þ should be 1:2. Ensure that carbonate is in high excess to
hydrogen chloride, because transferrin loading requires bicarbonate anions
(Richardson and Ponka, 1997). Incubate at room temperature overnight.
55Fe-loaded transferrin is separated from nonbound iron by desalting with
Hanks balanced salt solution (HBSS) on a PD-10 gel filtration column. The
elution process can be nicely followed because of the light brownish color
of iron-loaded transferrin. Dilute the eluate as desired, add 1% BSA, sterile
filter and store at 4 until use.
For loading with 55Fe, cells are seeded at low densities in complete
growth medium [e. g., after electroporation (see earlier)] and cultured for
3 or 4 days in the presence of 1 mM 55Fe-Tf. Because transferrin receptor
(TfR) expression, and consequently iron uptake, is cell density-dependent
(Ponka and Lok, 1999; Stehling et al., 2008), the comparison of different
samples requires identical growth conditions. The higher the degree of
confluence, the lower is the relevance of density for TfR expression.
Harvesting of cells by trypsinization removes surface-bound transferrin
and allows the determination of total cellular 55Fe uptake by scintillation
counting as described for yeast model systems (Stehling et al., 2007).
3.1.2. Preparation of membranes containing respiratory
chain complexes
Membrane preparations of 55Fe-loaded HeLa cells are used to visualize the
incorporation of radioactive iron into respiratory chain complexes by
BN-PAGE (Schägger, 2003). To prepare membrane samples, suspend
pelleted HeLa cells at an amount of 10 mg wet weight per 100 ml in BN
Mitobuffer (250 mM sucrose in 20 mM sodium phosphate, pH 7.0) and
homogenize cells with a tight-fitting, motorized glass/Teflon homogenizer
by 40 passes at a rotation of 2,000 rpm. Cell opening may be monitored by
trypan blue exclusion. Rinse the pestle with up to 1 ml BN Mitobuffer and
determine the total sample volume. Transfer the lysate in aliquots
corresponding to 20 mg HeLa cell wet weight into 1.50-ml microfuge
tubes and centrifuge at 13,000g for 10 min at 4 . After discarding the
supernatant, pellets may be used immediately or quick-frozen in liquid
nitrogen and stored at 80 until use.
For loading on a BN gel solubilize membrane preparations by suspending
each aliquot in 35 ml of BN gel solubilization buffer (50 mM NaCl, 50 mM
imidazole, 2 mM aminocaproic acid, 1 mM EDTA, pH 7.0). Suspending may
require mechanical dispersion (e.g., by a spatula). Add 10 ml of a 20% (w/v)
digitonin solution (dissolved in water at 95 ) and mix immediately by vortexing or pipetting. Clear suspension by centrifuging at 100,000g for 15 min at 4 .
Determine protein content of the clear supernatant to ensure equal loading of
the gel, add 10 ml of a 5% (w/v) Coomassie blue G250 solution (weighed in
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500 mM aminocaproic acid), vortex immediately, and finally add 5% (w/v)
glycerol. Samples with equal protein content (300 mg) are loaded on a BN
gel and electrophoresed. Inclusion of a respiratory complex-rich reference
sample into the same gel allows monitoring of separation and migration of
mammalian respiratory complexes (Fig. 12.3A) to verify the positions of the
respective HeLa cell complexes. The reference sample may be prepared from
bovine heart mitochondria (Schägger, 2003) and solubilized in digitonin/
Coomassie as described previously for the HeLa cell lysates.
3.1.3. Blue-native PAGE and autoradiography
Cast a 1.5-mm-thick BN gradient gel from 4 to 13% with a gradient mixer
and a peristaltic pump. The 4% BN gel mix (final volume approximately
35 ml) consists of 2.8 ml of a 49.5% acrylamide/3% bis-acrylamide solution
3
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Complex I
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NS
Complex II
A
B
C
D
Figure 12.3 Effects of huNfs1 or huInd1 depletion on respiratory complexes.
(A) Bovine heart mitochondria (BHM), solubilized in digitonin, resolved by BNPAGE, and stained with Coomassie blue G250. Note that supercomplexes containing
complex I, complex III dimer (III2), and varying amounts of complex IV are present
when digitonin is used. (B, C) HeLa cells were labeled with 55Fe-transferrin, fractionated, and mitochondrial membrane^containing samples were prepared. On
BN-PAGE, respiratory complex I is only detectable as part of a supercomplex and its
iron content decreases on application of the RNAi vectors huNfs1-R3 (B) and siIND1
(C). Complex II associated iron is only barely detectable in the huNfs1-deficient cells
(B). The most prominent band is that of ferritin [Ft], which partially fractionates into
the crude mitochondrial preparation. (D) In-gel complex I activity assay. Complex I
activity decreases on huNfs1 or huInd1 depletion by RNAi. Again, complex I is only
apparent as part of a supercomplex. A nonspecific band (NS) is apparent in the lower
part of the gel.
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(AA solution), 11.7 ml of a 3 BN gel buffer (75 mM imidazole in 1.5 M
aminocaproic acid), 20.5 ml water, 175 ml of a 10% (w/v) ammoniumperoxodisulfate solution (APS), and 30 ml N,N,N0 ,N0 -tetramethylethan-1,2diamine (TEMED). The 13% BN gel mix (final volume approximately
35 ml) is made up by 9.2 ml AA solution, 11.7 ml 3 BN gel buffer, 5.6 ml
glycerol, 8.5 ml water, 119 ml APS, and 11.9 ml TEMED. With the aid of a
long needle inserted to the bottom of the cast, we pour our gels from the
bottom up, the lower percentage acrylamide solution entering the cast first.
Adding 1 ml of water to the cast, before pouring the gel, forms a level top
edge to the separating gel. Adjust the flow rate to an intermediate speed
(4 ml/min) to allow for good gradient formation and to avoid premature
polymerization. An ambient temperature of 4 will facilitate casting. Polymerization is promoted by carefully transferring poured gels to room
temperature. When the separating gel has congealed, add a stacking gel
(final volume approximately 29 ml) consisting of 1.75 ml AA solution,
8.3 ml 3 BN gel buffer, 18.45 ml water, 200 ml APS, and 20 ml
TEMED. Gel run is initially performed with Deep Blue Cathode Buffer
(50 mM Tricine, 7.5 mM imidazole, 0.02% Coomassie G250, pH 7.0) in
the top reservoir and 25 mM imidazole as anode buffer (bottom reservoir) at
100 V and 20 mA in the cold for 15 to 25 min to allow the protein to run
into the gel. The voltage is then turned up to 500 V. Once the leading edge
of the samples has migrated one-third the length of the gel, the Deep Blue
Cathode Buffer is exchanged by B/10 Cathode Buffer (50 mM Tricine, 7.5
mM imidazole, 0.002% Coomassie G250, pH 7.0) to diminish the amount
of Coomassie within the gel. After the run is completed, bovine heart
mitochondrial complexes I to V of the reference sample are visible without
further staining (Fig. 12.3A); however’ Coomassie staining may be necessary to visualize all of the respiratory complexes. This can be achieved by
shaking the respective gel strip for 1 to 2 h in 0.025% Coomassie blue G250
dissolved in 10% acetic acid and then washing the gel in 10% acetic acid for a
few hours. The remaining gel containing the HeLa cell samples is dried
either by use of a gel dryer or gel drying film. Iron-containing complexes
are visualized by exposure to a phosphor storage screen and phosphorimaging (Fig. 12.3B,C), or by autoradiography. In the latter case, appropriate
signal enhancing systems and a longer exposure time are required.
3.2. Assessing the subunit composition of respiratory
complexes by two-dimensional BN-PAGE
Maturation of respiratory complexes occurs in multiple steps by which
individual subunits or preformed subcomplexes are successively assembled
to build the functional enzyme (Coenen et al., 2001; Fernandez-Vizarra
et al., 2007; Pickova et al., 2005; Vogel et al., 2007). In the absence of
general assembly factors like Nfs1 or specific assembly factors like Ind1,
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Fe/S subunits are lacking their cofactors and frequently become unstable.
As a consequence, assembly of the respective complexes is impaired, and
protein levels of individual subunits may decrease by proteolysis (Bych et al.,
2008; Gerbeth et al., manuscript in preparation; Sheftel et al., manuscript in
preparation). Subunit composition of respiratory complexes or preassembled subcomplexes can be assessed by 2-D BN-PAGE (Schägger, 2003).
Respiratory complexes are first separated by BN-PAGE, and then individual complexes and subcomplexes are deconstructed into their individual
subunits by SDS-PAGE in a second dimension.
To run the second dimension, soak 1.5-mm-thick BN gel strips containing mitochondrial membrane preparations resolved by BN-PAGE (see
section 3.1.) for 15 to 30 min in 1% (w/v) sodium dodecylsulfate (SDS) at
room temperature. Take care that the strips are completely covered. Place
each strip horizontally on a gel plate containing the ‘‘ears’’ and squeeze it by
assembly of the second plate with 0.7- to 1.0-mm spacers. Prepare a 10% 2D
SDS-gel solution consisting of 13.3 ml 2-D SDS-gel buffer (0.3% w/v SDS
in 3 M TRIS-HCl, pH 8.45), 8.13 ml AA solution (see section 3.1.3), 4 ml
glycerol, 14.4 ml water, 200 ml APS, and 10 ml TEMED. Pour the 2-D
SDS-gel solution nearly up to the gel strip and overlay with water. After
polymerization, push down the gel strip to improve contact to the SDS-gel,
remove the remaining water, and pour a 10% BN gel solution (1.04 ml AA
solution, 1.67 ml 3 BN gel buffer, 2.26 ml water, 28 ml APS, 2.8 ml
TEMED; c. f. section 3.1.3.) along sides but not on top of the gel strip. Run
the gel at room temperature for 4 to 5 h at 200 V (or 50 V overnight, not
exceeding 50 mA in each case) with 0.1 M TRIS, 0.1 M Tricine, and 0.1%
w/v SDS as 2-D cathode buffer and 0.2 M TRIS-HCl (pH 8.9) as anode
buffer. Resolved proteins can be detected within the gel by conventional
silver staining (Sheftel et al., manuscript in preparation). Alternately, individual proteins may be identified by immunoblotting subsequent to the
gel run.
4. Analysis of Respiratory Complex Function
4.1. Determination of lactate formation
Respiratory chain deficiencies are often associated with excess lactate formation. NADH generated by glycolysis and the citric acid cycle cannot be
oxidized by complex I, either because of a direct dysfunction of this enzyme
(Triepels et al., 2001) or because of an impaired electron drain caused by
dysfunctions downstream of the electron transport chain (Munnich and
Rustin, 2001). Instead, NADH is used by lactate dehydrogenase to convert
pyruvate to lactate. Increased lactate levels in the culture medium of HeLa
cells are thus an indication for impaired respiratory complex activity on
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huNfs1 and huInd1 deficiency. We use multiwell plates to facilitate the
biochemical determination of lactate levels in the respective tissue culture
supernatants. The assay is based on the reduction of NADþ because of the
action of lactate dehydrogenase that converts lactate to pyruvate (Gawehn,
1988). To shift the equilibrium of the reaction toward the conversion of
lactate, D-glutamate-pyruvate transaminase and D-glutamate are added to
transfer the amino group to pyruvate yielding D-alanine and 2-oxoglutarate.
To determine lactate concentrations, dilute conditioned tissue culture
supernatants 10-fold in water. Prepare aqueous lactate reference solutions
containing 0.1 to 1.5 mM lactate and 10% nonconditioned culture medium.
If the tissue culture cells produce large amounts of lactate, dilutions of the
media have to be increased respectively. Load 11.2 ml of each sample as well
as a solvent only blank of 10-fold diluted nonconditioned culture medium
into wells of a 96-well UV/Vis plate. Immediately before use, make up the
reaction buffer (pH 10.0) containing 250 mM glycylglycine, 500 mM
glutamate, 4.8 mM NADþ, and 40 U/ml D-glutamate-pyruvate transaminase (GPT). Pipette 238 ml of this reaction buffer to each sample and begin
measuring absorption at l ¼ 340 nm. Allow the reaction to proceed for
approximately 5 min until the signal has stabilized. Add 3 ml L-lactate
dehydrogenase solution (LDH, 16.5 U) to each well and resume data
acquisition for approximately 40 min. When other sample sizes are used,
volumes of buffers and reagents have to be adjusted accordingly. The
increase in the optical density at l ¼ 340 nm is calculated by subtracting
the absorbance before the addition of LDH from the absorbance plateau
reached at the end of the assay. The absolute amount of lactate in each tissue
culture sample can be deduced from the absorbance changes occurring in
the lactate reference samples.
4.2. Determination of complex I activity by in-gel
activity staining
Although increased lactate formation is a general indication for impaired
respiratory chain activity upon huNfs1 and huInd1 depletion, the direct
determination of the activities of individual respiratory complexes is
providing more quantitative information about the functional deficits. We
frequently apply 1-dimensional BN-PAGE to individually determine the
activity of complex I (Zerbetto et al., 1997). The assay is based on the
NADH-dependent reduction of p-nitrotetrazolium blue chloride (NBT),
which is leading to a discoloration and in-gel precipitation of the dye.
For activity staining, apply an adequate volume of 3 mM p-nitrotetrazolium blue chloride dissolved in 5 mM TRIS/HCl (pH 7.4) to sufficiently
cover the gel. Next, add NADH to a final concentration of 120 mM and
immediately immerse the gel in the reaction solution. Gently shake at room
temperature for 1 to 30 min, monitoring the gel for the formation of violet
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bands near the top of the gel (Fig. 12.3D; complex I monomer is approximately 1 MDa, whereas supercomplexes can be approximately twice this
size). When an appropriate time has been reached, decant the liquid and fix
the gel by adding 10% (v/v) acetic acid in 50% (v/v) methanol and shaking
for 1 to 2 h. If necessary, exchange the fixing solution as desired to remove
the residual blue dye from the gel, and then place the gel in water. Scan the
gel for quantification before drying.
4.3. Determination of complex I activity by
spectrophotometry
Despite that enzymatic in-gel activity can be densitometrically evaluated,
spectrophotometry allows a more quantitative determination of complex I
activity. Examining the oxidation of deaminodiphosphopyridine nucleotide
(dNADH), a NADH derivative, provides a direct evaluation of the enzymatic activity of this respiratory complex (Grgic et al., 2004). In reference
samples, nonspecific dNADH oxidation can be monitored by inhibition of
complex I by adding 2-decyl-4-quinazolinyl amine (DQA).
Prepare complex I reaction buffer (1.5 ml is required per sample) by
adding 2 mM KCN, 60 mM decylubiquinone (DBQ), and 100 mM dNADH
to an appropriate volume of complex I stock buffer consisting of 20 mM
sodium MOPS (pH 7.4), 50 mM NaCl, and 2.5 mg/ml BSA, and gently
invert several times to mix. After adding the DBQ to the reaction solution,
it becomes sensitive to air and must not be vigorously shaken. We find that
measurements are more stable when the reaction buffer is allowed to rest at
room temperature for approximately 20 min subsequent to mixing. Split the
reaction buffer in half, and to one half add the complex I inhibitor DQA to
yield 27 mM. Pipette each half of the reaction buffer (750 ml; DQA) into
one of two UV/Vis cuvettes. To each cuvette, add 30 to 50 mg protein from
the mitochondrial membrane preparations (see section 3.1.2.) and mix
samples with a plastic stirring spatula. Changes in absorbance are measured
for several minutes. Complex I activity is determined by the rate of
dNADH oxidation, which results from the difference in absorption at
340 nm between the two samples that lack or contain DQA. The molar
extinction coefficient for dNADH at l ¼ 340 nm is similar to NADH and
amounts to e340 nm ¼ 6200 M1 cm1.
4.4. Analysis of enzyme activities in multiwell plates
The continuous spectrophotometric measurement of enzyme activities in
individual cuvettes is time-consuming. Because the analysis of diverse
enzyme activities in differentially treated cells by replicative samples results
in an exponential increase in sample number, we perform parallel enzyme
assays in multiwell plates with a Tecan infinite M200 device (Tecan,
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Switzerland) in its kinetics mode. The plates are placed on a bottom-cooled
dry surface and samples are loaded as required. The amount of protein
deployed has to be adjusted so that low enzyme activities can be followed
for at least 10 to 15 min. Sample volumes should not exceed 10% of the final
reaction volumes in one well. Buffer stocks can be prepared in advance,
but enzyme substrates should be added fresh. Right before analysis, the
multiwell plate is placed at room temperature and equal volumes of prewarmed reaction buffer are poured quickly into the wells with a repeatvolume pipettor. The final volume within a well does not require further
adjustment, because the dilution of the reaction buffer by the protein
sample is not influencing enzyme-related changes in total optical density.
Although slightly increased final volume increases the light path, the simultaneously diminished concentration of the respective absorbing substance is
compensating the effects on extinction. Care has to be taken not to produce
air bubbles; otherwise, they have to be destroyed by tapping with a sharp
needle. Immediately after addition of reaction buffer, the multiwell plate is
placed into the reader device and orbitally shaken for 5 sec with an amplitude of 2.5 mm. Changes in optical densities are recorded at an ambient
temperature of 37 .
4.4.1. Determination of complex II (SDH) activity
Mammalian respiratory complex II (succinate: ubiquinone oxidoreductase) is
an interface between the citric acid cycle and the respiratory chain (Cecchini,
2003). It converts succinate to fumarate, thereby feeding electrons to the
quinone pool of the electron transport chain. Complex II consists of four
subunits, with two of them, SdhA and SdhB, exposed to the mitochondrial
matrix, whereas subunits SdhC and SdhD serve as membrane anchors.
Because of its enzymatic activity, complex II is also referred to as succinate
dehydrogenase (SDH). Three Fe/S clusters are located in SdhB, channeling
electrons from FAD in SdhA to redox centers in the membrane domain. The
consequence especially of huNfs1 deficiency on SDH activity can easily be
tested in total HeLa cell samples with a multiwell plate-based, enzymatic assay
(Hatefi and Stiggall, 1978). The method is based on the oxidation of succinate
to fumarate with concomitant reduction of the blue dye 2,6-dichloro-N-(4hydroxyphenyl)-1,4-benzoquinoneimine (DCPIP), rendering it colorless, by
use of decylubiquinone as the electron carrier.
To determine SDH activity, load equal amounts of HeLa cell samples
corresponding to 4 to 12 mg of total protein in each of two wells of a 48-well
plate. Prepare SDH reaction buffer by adding 1 mM KCN, 60 mM DBQ,
and 7,5 mM disodium succinate solution to a SDH stock buffer consisting of
50 mM TRIS/SO4 (pH 7.4), 0.1 mM EDTA, 70 mM DCPIP, and 0.1%
Triton X-100. Subsequently, dispense 750 ml of this SDH reaction buffer in
one of the two sample wells. Because DBQ may transfer electrons derived
from other sources than SDH, nonspecific DCPIP reduction can be
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determined by specific inhibition of SDH with malonate. Thus, immediately add 12 mM disodium malonate solution to the remaining SDH
reaction buffer, dispense 750 ml of this reference buffer into the remaining
second sample well, and start measurement. When multiwell plates of other
sizes are used, protein amounts and buffer volumes have to be adjusted
accordingly. DCPIP reduction will proceed for 20 to 30 min. The molar
extinction coefficient of DCPIP is e600 nm ¼ 21,000 M1 cm1. SDH
activity is determined by calculating the difference between total and
nonspecific DCPIP bleaching.
4.4.2. Determination of complex IV (COX) activity
Respiratory complex IV (cytochrome c oxidase, COX) is the final member
of the electron transport chain, transferring electrons from cytochrome c to
molecular oxygen (Ludwig et al., 2001). It consists of 13 subunits and
contains two heme and two copper centers in its functional core that is
formed by three mitochondria-encoded polypeptide chains. Because Fe/S
clusters are absent, the enzyme is often used as internal reference for the
analysis of mitochondrial Fe/S proteins. However, COX activity is not
inevitably invariable but regulated by various physiologic factors such as the
intramitochondrial ATP/ADP-ratio, cAMP and calcium-dependent phosphorylation and dephosphorylation, and subunit expression (Fontanesi
et al., 2008; Ludwig et al., 2001). In HeLa cells, we observed a cell-density
dependent increase in COX activity (unpublished). Moreover, depletion of
general assembly factors for respiratory complexes like Nfs1, as well as other
members of the ISC assembly machinery, impairs cellular iron homeostasis
(Biederbick et al., 2006; Rouault and Tong, 2008) with a potential impact
on heme formation and COX activity. Thus, use of COX for standardizing
activities of other mitochondrial enzymes is not unequivocal. In any case,
we suggest determination of COX activity in cultured cells grown at several
cell densities to obtain reference values. Measurements can be done with
total HeLa cell lysates by examining COX-mediated oxidation of cytochrome c with a multiwell plate-based enzymatic assay (Birch-Machin and
Turnbull, 2001; Trounce et al., 1996).
4.4.2.1. Preparing reduced cytochrome c Before its use as COX substrate, cytochrome c (cyt-c) has to be reduced. Dissolve the oxidized protein
at a concentration of 100 mg/ml in 30 mM potassium phosphate buffer (pH
7.2) and add an equal volume of 20 mM sodium dithionite. The color of the
solution will change from a dark brownish red to a light red. Desalt the
reduced cyt-c anaerobically by passage through a PD-10 column with 15
mM potassium phosphate buffer (pH 7.2). Reduction and concentration of
the eluted protein can be determined spectrophotometrically. The molar
extinction coefficient is e550 nm ¼ 19,500 M1 cm1. Quick-freeze
appropriate aliquots in liquid nitrogen and store at 80 until use.
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4.4.2.2. Measuring COX activity Load HeLa cell samples corresponding
to 0.5 to 2 mg of total protein in a 96-well plate. Immediately before use, add
20 mM reduced cyt-c to the COX stock buffer (15 mM potassium phosphate,
0.1% BSA, 0.5 mM dodecylmaltoside, pH 7.2) and dispense 250 ml of this
COX assay buffer to each protein sample. Because cyt-c is rapidly oxidized
in air, we usually load another three wells with COX assay buffer alone as an
internal standard to determine nonspecific changes in cyt-c absorbance.
Alternately, COX assay buffer containing 1 mM KCN may be dispensed
to replicate samples of HeLa cells as a reference, but in our hands cyt-c is
oxidized as fast as in the standard wells mentioned previously. Immediately
place the multiwell plate into the reader device, because cyt-c oxidation will
proceed only for 10 to 15 min. COX activity is determined by calculating
the difference between total and nonspecific cyt-c oxidation. The molar
extinction coefficient of reduced cyt-c is l550 nm ¼ 19,500 M1 cm1.
4.4.3. Determination of citrate synthase activity
Because of the potential variability of COX activity, we instead prefer citrate
synthase (CS) as an alternative mitochondrial reference enzyme to scrutinize
mitochondrial function. Although CS is not a direct member of the respiratory chain, it is indirectly linked by means of its participation in the citric
acid cycle to produce the substrates for respiratory complexes I and II. CS is
a homodimer, expediently not containing any cofactor, resides within the
mitochondrial matrix, and catalyzes the condensation of acetyl-coenzyme A
with oxaloacetate to form citrate (Wiegand and Remington, 1986). In our
experience, CS activity did not significantly change by any manipulation we
performed on mitochondria of HeLa cells. Enzymatic activity can easily be
measured in total HeLa cell samples with a sensitive multiwell plate-based
enzymatic assay according to a published procedure (Srere et al., 1963). The
method builds on the hydrolytic cleavage of 5,50 -dithiobis[2-nitrobenzoic
acid] (DTNB, Ellman’s reagent) to form 5-mercapto-2-nitrobenzoate by
free sulfhydryl groups of coenzyme A that is liberated on the condensation
of acetyl-CoA with oxaloacetate.
To determine CS activity, load equal amounts of HeLa cell samples (2 to
5 mg of total protein) in each of two wells of a 48-well plate. Prepare CS
reference buffer by adding 175 mM acetyl-CoA to the CS stock buffer
(50 mM TRIS/HCl, 100 mM NaCl, 0.5 mM DTNB, 0.1% Triton
X-100, pH 8.0) and dispense 750 ml in one of the two wells to monitor
nonspecific DTNB hydrolysis. Quickly add 1 mM sodium oxaloacetate to
the reference buffer to obtain complete CS assay buffer, dispense 750 ml to
the second HeLa cell sample, and start measurement. When multiwell plates
of other sizes are used, protein amounts and buffer volumes have to be
adjusted accordingly. DTNB hydrolysis may proceed for more than 30 min,
depending on the protein amount applied. Take care to not load too
much protein to avoid depletion of the substrates. The molar extinction
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coefficient of the DTNB hydrolysis product 5-mercapto-2-nitrobenzoate is
e412 nm ¼ 13,300 M1 cm1. CS activity is determined from the
difference between total and nonspecific DTNB hydrolysis.
5. Concluding Remarks
Maturation of respiratory chain complexes requires the concerted
interaction of multitudinous assembly factors that either directly assist complex formation or act indirectly by means of their involvement in cofactor
synthesis and/or insertion. The intricate dependence of the respiratory
chain on Fe/S clusters makes complexes I, II, and III excellent targets for
the molecular and functional analysis of Fe/S protein biogenesis. Thus,
examination of the catalytic activities and the Fe/S status of individual
respiratory complexes considerably contributes to our understanding of
how Fe/S cluster are synthesized and inserted into apoproteins. Vice
versa, studying Fe/S protein formation may facilitate the identification of
new assembly factors, such as huInd1 for respiratory complex I. The
experimental strategies and techniques described herein may help in future
investigations to deepen our insights into both general mechanisms of the
assembly of cellular Fe/S proteins and the specific maturation pathways of
respiratory chain complexes.
ACKNOWLEDGMENTS
We acknowledge the generous gift of DQA by Dr. Hermann Schägger as well as his
invaluable advice and the technical assistance of Christian Bach. We also thank Carolin
Gerbeth for providing the huNfs1 immunoblot. A. D. S. is supported by fellowships from
the Fonds de la Recherche en Santé Québec and the Alexander-von-Humboldt Foundation.
R. L. acknowledges support from Deutsche Forschungsgemeinschaft (SFB 593 and TR1,
Gottfried-Wilhelm Leibniz program, and GRK 1216), the German-Israeli Foundation GIF,
Rhön Klinikum AG, von Behring-Röntgen Stiftung, and Fonds der chemischen Industrie.
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